This Article Abstract Full Text (PDF) Alert me when this article is cited Alert me if a correction is posted Citation Map Services Email this article to a friend Similar articles in this journal Similar articles in PubMed Alert me to new issues of the journal Download to citation manager Request Permissions Citing Articles Citing Articles via HighWire Citing Articles via Google Scholar Google Scholar Articles by Aikawa, E. Articles by Schoen, F. J. Search for Related Content PubMed PubMed Citation Articles by Aikawa, E. Articles by Schoen, F. J. Related Collections Valvular heart disease CV surgery: valvular disease

(Circulation. 2006;113:1344-1352.)
© 2006 American Heart Association, Inc.

Valvular Heart Disease

Human Semilunar Cardiac Valve Remodeling by Activated Cells From Fetus to Adult

Implications for Postnatal Adaptation, Pathology, and Tissue Engineering

Elena Aikawa, MD, PhD; Peter Whittaker, PhD; Mark Farber, MS; Karen Mendelson, MS; Robert F. Padera, MD, PhD; Masanori Aikawa, MD, PhD; Frederick J. Schoen, MD, PhD

From the Department of Pathology (E.A., M.F., K.M., R.F.P., F.J.S.) and Cardiovascular Division, Department of Medicine (M.A.), Brigham and Women’s Hospital, Harvard Medical School, Boston, Mass; Departments of Emergency Medicine and Anesthesiology (P.W.), University of Massachusetts Medical School, Worcester, Mass; and Center for Molecular Imaging Research (E.A.), Massachusetts General Hospital, Boston, Mass.

Correspondence to Frederick J. Schoen, MD, PhD, Department of Pathology, Brigham and Women’s Hospital, 75 Francis St, Boston, MA 02115 (e-mail fschoen{at}partners.org), or Elena Aikawa, MD, PhD, Center for Molecular Imaging Research, Massachusetts General Hospital, 149 13th St, Charlestown, MA 02129 (e-mail eaikawa@partners.org).

Received September 27, 2005; revision received December 24, 2005; accepted January 9, 2006.

	Abstract

Top Abstract Introduction Methods Results Discussion References

 
Background— The evolution of cell phenotypes and matrix architecture in cardiac valves during fetal maturation and postnatal adaptation through senescence remains unexplored.

Methods and Results— We hypothesized that valvular interstitial (VIC) and endothelial cell (VEC) phenotypes, critical for maintaining valve function, change throughout life in response to environmental stimuli. We performed quantitative histological assessment of 91 human semilunar valves obtained from fetuses at 14 to 19 and 20 to 39 weeks’ gestation; neonates minutes to 30 days old; children aged 2 to 16 years; and adults. A trilaminar architecture appeared by 36 weeks of gestation but remained rudimentary compared with that of adult valves. VECs expressed an activated phenotype throughout fetal development. VIC density, proliferation, and apoptosis were significantly higher in fetal than adult valves. Pulmonary and aortic fetal VICs showed an activated myofibroblast-like phenotype (-actin expression), abundant embryonic myosin, and matrix metalloproteinase-collagenases, which indicates an immature/activated phenotype engaged in matrix remodeling versus a quiescent fibroblast-like phenotype in adults. At birth, the abrupt change from fetal to neonatal circulation was associated with a greater number of -actin–positive VICs in neonatal aortic versus pulmonary valves. Collagen content increased from early to late fetal stages but was subsequently unchanged, whereas elastin significantly increased postnatally. Collagen fiber color analysis revealed a progressive temporal decrease in thin fibers and a corresponding increase in thick fibers. Additionally, collagen fibers were more aligned in adult than fetal valves.

Conclusions— Fetal valves possess a dynamic/adaptive structure and contain cells with an activated/immature phenotype. During postnatal life, activated cells gradually become quiescent, whereas collagen matures, which suggests a progressive, environmentally mediated adaptation.

Key Words: cells • collagen • remodeling • tissue engineering • valves

	Introduction

Top Abstract Introduction Methods Results Discussion References

 
An understanding of extracellular matrix (ECM) architecture and cellular changes that occur in cardiac valves during fetal development, maturation, and aging would provide mechanistic insights into the pathogenesis of congenital and acquired valve abnormalities, aid assessment of therapeutic strategies for valve disease, and assist in the development of regenerative and tissue-engineered approaches to valve repair and replacement. Although the mechanisms of valvulogenesis in early cardiac development have been elucidated,^1–5 subsequent cellular and ECM maturation, remodeling, and growth processes, particularly in humans, remain unexplored.

ECM in general and collagen in particular play a crucial role in valve function and durability. Indeed, valves must accommodate substantial changes in hemodynamic environment and structure throughout a lifetime. Furthermore, valvular interstitial (VIC) and endothelial cell (VEC) functions likely influence ECM synthesis and remodeling. For example, activated myofibroblast-like VICs are critical to collagen metabolism and are altered in some valve diseases, including myxomatous degeneration of the mitral valve.⁶ We recently showed that large populations of VICs undergo phenotypic modulation to become activated myofibroblasts and return to quiescent fibroblasts during adaptive remodeling in response to changing environmental conditions, which we have observed in long-term pulmonary autografts and in tissue-engineered valves.^7–9 We therefore tested the hypothesis that phenotypic changes in valvular cells, determined by altered expression of cytoskeletal and surface proteins and by proteolytic enzymes, regulate age-associated structural remodeling and adaptation of the ECM. Specifically, we characterized phenotypes and turnover of VIC and VEC in human fetal, neonatal, child, and adult semilunar valves and correlated cellular changes with alterations in valvular ECM.

	Methods

Top Abstract Introduction Methods Results Discussion References

 
Morphological Characterization
We studied 91 human semilunar valves: fetal valves in the second trimester (14 to 19 weeks’ gestation; n=11) and third trimester (20 to 39 weeks’ gestation; aortic n=10, pulmonary n=10); neonatal (early neonatal with minutes to hours of survival: aortic n=5, pulmonary n=5; late neonatal [up to 30 days of survival]: aortic n=5, pulmonary n=5); valves from children (aged 6.0±1.6 years; aortic n=10, pulmonary n=10); and valves from normal adults (aged 50.1±2.5 years; aortic n=10, pulmonary n=10) obtained at autopsy according to a protocol for tissue use approved by the Human Research Committee at Brigham and Women’s Hospital. No patient had documented cardiac disease or conditions known to predispose to heart valve disease. Examination revealed no evidence of valve disease, and the fetal heart morphology appeared appropriate for the gestational age.

Valves were fixed in 10% buffered formalin, cut radially in the central cusp region through the adjacent arterial wall, and embedded in paraffin. Sections cut at 6-µm thickness were stained with hematoxylin and eosin for general morphological assessment. We used Movat pentachrome stain to differentiate connective tissue elements (collagen, elastin, and proteoglycans) and picrosirius red viewed with circularly polarized light to assess collagen architecture.^6,10 All 3 cusps from each aortic and pulmonary valve were analyzed.

Characterization of Cell Phenotype and Turnover
Markers of cell phenotype and synthetic products were identified by immunohistochemistry. We defined myofibroblasts like VICs as cells that had antibody reaction to -smooth muscle actin (-SMA, 1A4, Dako, Carpenteria, Calif).^6,11 Mouse monoclonal antibodies against human metalloproteinase-1 (MMP-1/collagenase-1) and MMP-13/collagenase-3 (Calbiochem, San Diego, Calif) were used to determine proteolytic enzyme expression. Cell activation was also demonstrated by a monoclonal antibody against human SMemb¹² (a nonmuscle myosin also known as MHC-B, expressed by embryonic or activated mesenchymal cells). Phenotypic changes in valvular endothelium were assessed by antibodies to CD31 (Dako), intercellular adhesion molecule-1 (ICAM-1; US Biological, Swampscott, Mass), and vascular cell adhesion molecule-1 (VCAM-1; US Biological). Immunohistochemistry methods used avidin-biotin-peroxidase after antigen unmasking pretreatment with proteinase K for 5 minutes at room temperature. Adjacent sections treated with nonimmune IgG provided controls for antibody specificity. Cell proliferation was demonstrated with the Ki-67 (MIB-1, Dako) antigen, which is present during active phases of the cell cycle but absent in resting cells, and cell apoptosis was demonstrated by terminal dUTP nick end-labeling (TUNEL) assay (Intergen, New York, NY). Commercially available human lymph node and thymus sections provided positive controls. Images were captured and analyzed with a digital camera (Nikon DXM1200-F, Nikon Inc, Melville, NY) and imaging software (ACT-1 version 2.63, Nikon).

Cell density was measured as mean number of cells per 10 high-power fields (x400) and expressed as cell number per square millimeter of the tissue section in each cusp. Cell proliferation and apoptosis indices were calculated as the ratio of Ki-67 and TUNEL-positive cells, respectively, to the total number of valve cusp cell nuclei. The ratio of -SMA, MMP-1, MMP-13, and SMemb-positive cells to the total cell number was averaged over 5 representative high-power fields. The ratio of ICAM-1– and VCAM-1–positive cells to total CD31-positive cells was calculated for each cusp. Only cells expressing an antigen of interest (defined as red reaction product associated with blue hematoxylin nuclear counterstain) were counted. All measurements were done by 2 independent researchers.

Collagen and Elastin Assessment
We used picrosirius red staining in conjunction with polarized light microscopy to assess collagen fiber content, thickness, and organization and Movat pentachrome to detect elastin (Olympus BX51 microscope, Olympus America Inc, Melville, NY).

Collagen Content and Fiber Color
With this microscopy combination, collagen fiber color is determined by thickness; the color changes from green to yellow to orange to red as thickness increases.¹³ We assessed the proportion of different colored fibers using published methods.^14,15 Briefly, we recorded images viewed with circularly polarized light¹⁶ using a digital camera (DP11, Olympus) and analyzed 2 regions (260x200 µm) per valve. Images were resolved, using an automated software function (SigmaScan Pro; SPSS Inc, Chicago, Ill), into their hue, saturation, and value components. We retained only the hue component, and a histogram of hue frequency was obtained from the 8-bit hue images that contained 256 possible colors, defined as follows; red 2 to 9 and 230 to 256, orange 10 to 38, yellow 39 to 51, and green 52 to 128.¹⁴ All other hue values corresponded to interstitial space, confirmed by inspection. We determined the number of pixels within each hue range and expressed this as a percentage of the total number of collagen pixels, which in turn was expressed as a percentage of the total number of pixels in the regions analyzed to give collagen content.

Collagen Orientation
The optical properties of birefringent materials, such as collagen, can be exploited to determine their 2D orientation.¹⁷ We used a recently developed automated system (PolScope, CRI, Inc, Woburn, Mass) to measure collagen fiber orientation at 100 locations in each valve, with locations selected by construction of a 10x5-point rectangular grid overlying 2 regions in each valve. An orientation frequency distribution was obtained for each region. Briefly, the system uses an electronically controlled liquid crystal compensator to acquire and record images, with a CCD camera, at 4 predetermined compensator settings. Algorithms are used to calculate the orientation angle of the slow optical axis of the birefringent collagen from the data obtained in these images¹⁸ given the property that the slow optical axis of collagen corresponds to the long-axis orientation of the fibers. Each distribution was analyzed with circular statistics, which are methods designed to examine directional data.¹⁹ The degree of fiber alignment was assessed by calculation of the angular deviation of the distribution, the circular statistics’ equivalent of standard deviation; the smaller the angular deviation, the more aligned the fibers.¹⁷

Elastin Content
Movat pentachrome stains elastin black, collagen yellow, and glycosaminoglycans blue-green. Elastin-positive areas were measured in 3 high-power fields of each cusp in the ventricularis layer with imaging software (IPLab version 3.9.3; Scanalytics, Inc, Rockville, Md).

Statistical Analyses
Data are presented as mean±SEM. Group differences were evaluated with ANOVA followed by pairwise comparisons with the Tukey-Kramer post hoc test; however, t tests were used when 2 groups were compared. Probability values less than 0.05 were considered significant.

The authors had full access to the data and take full responsibility for its integrity. All authors have read and agree to the manuscript as written.

	Results

Top Abstract Introduction Methods Results Discussion References

 
Fetal Valves Have an Incomplete Morphological Differentiation
At 14 weeks of gestational age, the ECM of fetal valves was predominantly composed of glycosaminoglycans, as indicated by the strong blue-green staining with Movat pentachrome (Figure 1). The overall valve architectural structure was homogeneous and lacked distinguishable layers. By 20 weeks, fetal valves had a bilaminar structure with sparse, loose, and unorganized collagen. A trilaminar structure containing elastin in the ventricularis and increased collagen in the fibrosa became apparent by 36 weeks of gestation but remained incomplete versus the normal adult valve structure²⁰ (Figure 1).

View larger version (68K): [in this window] [in a new window]   Figure 1. ECM composition of human cardiac valves from fetus to adult. At 14 to 19 weeks, fetal valves were composed mostly of glycosaminoglycans with low elastin and collagen. At 20 to 39 weeks, fetal valves had a bilaminar structure that contained elastin in the ventricularis and increased unorganized collagen in the fibrosa. A trilaminar structure became apparent in children’s valves but remained incomplete compared with normal adult valve layered architecture with collagen in the fibrosa, glycosaminoglycans in the spongiosa, and elastin in the ventricularis. Top, Movat pentachrome; bottom, picrosirius red under circular polarized light. Original magnification x200.

Fetal Valves Have High Cell Density and Proliferation Index
We found a progressive decrease in VIC density from the second and third trimesters to children’s and adult valves (3032±190, 2629±214, 1410±100, and 351±58 cells per mm², respectively; P<0.001 versus adults; Figure 2A). Nuclei that stained positively for Ki-67, an index of cell proliferation, were distributed diffusely throughout the cusp at 14 to 19 weeks’ gestation, whereas stained cells at 20 to 39 weeks of gestation and in children’s valves appeared predominantly near the arterial surface (Figure 2B). Fetal valves at 14 to 19 weeks’ gestation had higher VIC proliferation indices (26.4±1.3%, P<0.001 versus other groups), with an approximate 80% reduction in the third trimester (6.3±0.7%) and in children (4.2±0.5%) and a >90% reduction in adult valves (1.7±0.6%; Figure 2C). Apoptosis continued at a low rate throughout life (0.5±0.1%, 3.3±0.3%, 1.1±0.1%, and 0.5±0.1%, respectively; P<0.001 for late fetal valves versus other groups). Of note, fetal valves at 20 to 39 weeks of gestation contained more VICs that were undergoing apoptosis, predominantly located near the arterial surface (Figure 2C). The ratio of proliferation to apoptosis was high in fetal valves at 14 to 19 weeks (52.8; P<0.05) but was considerably lower in late gestation and in children and adults (1.9, 3.8, and 3.4, respectively).

View larger version (38K): [in this window] [in a new window]   Figure 2. Interstitial cell turnover in fetal, child, and adult valves. A, Progressive decrease in VIC density throughout life. *Significant differences in cell density between groups. B, Nuclei staining positively for Ki-67 (MIB-1) were distributed diffusely through the cusp at 14 to 19 weeks of gestation, whereas Ki-67 and TUNEL-positive cells at 20 to 39 weeks appeared predominantly near the arterial surface. Note that VICs in adult valves showed undetectable levels of proliferating and apoptotic cells. Bar=50 µm. Magnification x400. C, Fetal valves at 14 to 19 weeks of gestation had higher VIC proliferation indices, with >90% reduction in adult valves. *Significant differences in cell proliferation between groups. Apoptosis continued at a low rate throughout life. #Significant differences in cell death between groups. y.o. indicates years old.

Fetal Valves Contain Activated Cells That Overexpress Proteolytic Enzymes
In fetal valves at 14 to 19 weeks of gestation, VICs expressed the -SMA–positive phenotype, attributed to myofibroblasts, distributed predominantly in a subendocardial location below the outflow surface and throughout the deeper portion of the cusp. In contrast, -SMA–positive cells in adult valves were located predominantly (and exclusively) in regions adjacent to the endothelium (Figure 3A). We found larger myofibroblast-like VIC populations at 14 to 19 weeks than at 20 to 39 weeks’ gestation or in neonatal, child, or adult valves (28.3±1.6%, 17.4±1.3%, 20.5±2.2%, 6.2±1.7%, and 2.5±0.4%, respectively; P<0.001 versus fetal; Figure 3B). VICs characterized as activated/immature myofibroblasts expressed high levels of embryonic myosin heavy chain/SMemb and proteolytic enzymes (MMP-1, MMP-13), distributed in a homogeneous fashion throughout the cusp of fetal but not adult valves (Figure 3A; MMP-13 not shown). Fetal VICs at 14 to 19 weeks and 20 to 39 weeks and children’s VICs had larger numbers of SMemb-, MMP-1–, and MMP-13–positive cells versus the negligible numbers of VICs that expressed these proteins in adult valves (SMemb 80.9±3.1%, 75.2±4.1%, and 38±4.5% versus 7±1.0%; MMP-1 45.7±5.9%, 65.9±6.7%, and 61±5.6% versus 5.5±0.7%; MMP-13 18.5±2.5%, 47.3±4.8%, and 26.0±3.1% versus 3.5±0.7%, respectively; P<0.001 versus adult; Figure 3B).

View larger version (67K): [in this window] [in a new window]   Figure 3. Activated interstitial cells undergo evolution to a more quiescent phenotype during postnatal life. A, Larger VIC populations at 14 to 19 weeks expressed the -SMA–positive phenotype, attributed to myofibroblasts and high levels of SMemb and MMP-1, markers of activation, compared with negligible expression of these proteins in adult valves. Bar=50 µm. Magnification x400. B, Percent of interstitial cells positive for marker. *Significant differences in protein expression in adult valves. y.o. indicates years old.

At birth, after separation of the single fetal circulation into isolated pulmonary and systemic circulations, pulmonary arterial pressure decreases and aortic pressure increases.²¹ These hemodynamic changes were associated with decreased VIC activation in late neonatal (1 to 30 days of survival) pulmonary valves (6.3±0.8%; n=5), whereas aortic valve VICs remained activated (20.5±2.2%, n=5; P<0.05; Figure 4). Nevertheless, this change did not occur immediately, because the numbers of activated VICs in early neonatal valves with minutes to hours of survival were identical in both pulmonary (20.5%; n=5) and aortic valves (20.5%; n=5). Activated VICs continued to decrease after adaptation, because valves from individuals aged 2 to 16 years had similar numbers of activated VICs in pulmonary (6%; n=10) and aortic (6%; n=10) valves (Figure 4). Fetal VECs in the second and third trimesters and children’s valves consistently expressed the cell adhesion molecules ICAM-1 (99.0±7.2%, 98.7±4.6%, and 47.6±9.5% versus 5.0±0.5%) and VCAM-1 (54.0±9.4%, 68.2±13.9%, and 16.0±3.7% versus 0%), as well as SMemb (99.2±1.5%, 99.3±0.5%, and 71.3±10.3% versus 1.0±0.5%), MMP-1 (99.3±0.8%, 97.0±1.3%, and 71.7±9.1% versus 1.0±0.5%), and MMP-13 (98.4±2.5%, 95.3±1.7%, and 43.1±7.0% versus 0.9±0.3%), whereas normal adult VECs exhibited negligible expression levels of these proteins (P<0.001 fetal versus adult; Figure 5A and 5B).

View larger version (14K): [in this window] [in a new window]   Figure 4. Changes in VIC activation at birth. Abrupt hemodynamic changes at birth were associated with decreased VIC activation in late neonatal (days of survival) pulmonary valves (PV), whereas aortic valve (AV) VICs remained activated (P<0.05). Numbers of activated VICs in early neonatal valves (minutes to hours of survival) were identical in both pulmonary and aortic valves, which suggests that this change did not occur immediately.

View larger version (69K): [in this window] [in a new window]   Figure 5. Endothelial cells of fetal valves have an immature/activated phenotype. A, CD31-positive fetal VECs in the second and third trimesters and VECs in children’s valves consistently expressed SMemb, MMP-1, MMP-13, and cell adhesion molecules ICAM-1 and VCAM-1, whereas normal adult VECs mostly had negligible expression levels of these proteins. Bar=50 µm. Magnification x400. B, Percent of endothelial cells positive for maker. *Significant differences in protein expression in adult valves (P<0.001).

Collagen Content Plateaus In Utero, Whereas Elastin Content and Collagen Orientation Progressively Increase With Valve Maturity
Total collagen content, determined by picrosirius red–positive area, increased between the second (45.3±5.8%) and third (74.2±3.4%; P<0.001) trimesters but did not increase thereafter (Figure 6A). The progression from second to third trimesters was also associated with fiber color changes; the proportion of thin/green fibers decreased from 72.3±6.3% to 23.6±3.1% (P<0.001), whereas the proportion of thick/orange fibers increased from 5.2±1.6% to 29.5±7.0% (Figure 6A; P<0.05). We found no color difference between late fetal and children’s valves; however, the progression to adult valves saw further reductions in green fibers (7.1±1.7%) and increases in orange fibers (46.8±6.7%; P<0.05 versus child), even though there was no increase in total collagen. Although we found no intergroup difference in the proportion of red collagen pixels (range 0.9% to 1.8%; P=NS), our results indicate changes in collagen fiber thickness during valve maturation (Figure 6B).

View larger version (20K): [in this window] [in a new window]   Figure 6. Quantitative assessment of collagen content and orientation in cardiac valves from fetus to adult. A, Total collagen content increased between early and late fetal (P<0.001) stages but did not increase in children’s and adult valves. The progression from early to late fetal was also associated with collagen fiber thickness changes: the proportion of thin/green fibers decreased (P<0.001), whereas the proportion of thick/orange fibers increased (P<0.05). Note no difference between late fetal and children’s valves; however, the progression to adult valves saw further reductions in thin/green fibers and increases in thick/orange fibers (P<0.05 vs children), even though there was no increase in total collagen. B, Changes in collagen fiber thickness during valve maturation demonstrated by hue analysis. Color threshold images show the spatial distribution of green pixels, an indication of thin fibers, and orange pixels, an indication of thick, more mature fibers. Fetal valves contained more green collagen, whereas adult valves had more orange collagen. C, Angular deviation in adult valves significantly decreased (*P<0.01 vs other groups), which indicates increased collagen fiber alignment. D, Representative collagen fiber orientation distributions in early fetal and adult valves. *, #, indicate significant differences.

Although there was a small postnatal reduction in angular deviation, this change was not significant (Figure 6C). In contrast, angular deviation in adult valves decreased (P<0.01 versus other groups), which indicates a significant increase in fiber alignment, illustrated in the representative orientation distributions. These distributions represent the relative orientation of fibers within the region analyzed, rather than orientation relative to a specific direction (Figure 6D).

Elastin was measured as a black-positive area with Movat pentachrome stain and expressed as a percent of the area of the ventricularis layer. Elastin content was negligible at 14 weeks’ gestation (0.7±0.1%). The first discernable fibers appeared on the ventricular aspect in the base of the cusps at the end of the third trimester (3.8±0.5%). Total elastin content increased in children’s valves (8.2±2.2%) and was higher yet in adult semilunar valves (25.1±3.9%; P<0.05; Figure 7A). Adult aortic valves had more elastin (32.3±2.7%) than pulmonary valves (17.9±2.7%, P<0.001; Figure 7B).

View larger version (57K): [in this window] [in a new window]   Figure 7. Quantitative assessment of elastin content in cardiac valves. A, Elastin content was low in fetal valves. Total elastin content increased in adolescent valves and reached a maximum in adults. *Significant differences in elastin content (P<0.001); no difference was found between early and late fetal valves. B, Adult aortic valves had more elastin (black) than pulmonary valves. Inflow surface at upper right. Movat pentachrome. Magnification x200.

	Discussion

Top Abstract Introduction Methods Results Discussion References

 
This study focused on the evolution in cell phenotypes and ECM remodeling with valve maturation. We demonstrated that fetal valves possess a dynamic/adaptive structure and contain cells with an activated/immature phenotype. During postnatal life, activated cells gradually become quiescent, whereas collagen matures through increased fiber thickness and alignment, which suggests a progressive, environmentally mediated adaptation.

Semilunar Valve Architecture Changes Throughout Development and Into the Postnatal Period
The fetal circulation differs from that of adults owing to shunts that equalize left and right ventricular pressures: the foramen ovale between the right and left atrium and ductus arteriosus between pulmonary artery and aorta. Thus, fetal semilunar valves and the aorta and pulmonary trunk are subjected to equal pressures, which progressively increase during gestation.²¹

The present study revealed that fetal, second-trimester semilunar (aortic and pulmonary) valves lacked distinguishable layers, were composed primarily of proteoglycans, had no detectable elastin, had small amounts of disorganized collagen, and were histologically identical (likely because of a similar physiological environment during development²²). Valvular ECM elements accommodate repetitive shape and dimension changes throughout the cardiac cycle and must adapt to different mechanical stresses throughout life. Therefore, it is not surprising that pressure increases during gestation increased collagen and elastic fiber content and increased collagen fiber alignment (Figures 1 and 6). Nevertheless, fetal valve structure differed, even late in gestation, from that of adult valves, which have a trilayered architecture with a highly specialized and functionally adapted ECM (Figure 1). These observations suggest that valvular cusp morphogenesis continues throughout development and into the postnatal period. After birth, the foramen ovale closes and the ductus arteriosus occludes, which results in separation of the pulmonary and systemic circulations. Lower pressure in the pulmonary circulation is eventually reflected in adult pulmonary valve cusps that are more delicate than the aortic, whereas higher pressure in the systemic circulation produces thicker aortic cusps,⁷ with dense collagen fibers arranged parallel to the free edges of the cusp, predominantly on the arterial aspect, and more elastic fibers on the ventricular side (Figure 7B).

Turnover of Valve Interstitial Cells Is High During Fetal Development and Continues at a Low Rate Postnatally
During valve morphogenesis and development, cells of the cardiovascular system differentiate and proliferate and can migrate over large distances while they produce, degrade, and remodel ECM. Cells originating in the neutral crest, for example, migrate to the cardiovascular system and are critical to the formation of several structures, including the semilunar valves.²³ During valve formation, cardiac cushions appear as localized expansions of ECM, known as cardiac jelly. Endothelial cells then invade the cushions, transform into mesenchymal cells,¹ and eventually form the inflow (mitral and tricuspid) and outflow (aortic and pulmonary) valves, a process aided in part by cell proliferation²⁴ and apoptosis.²⁵ The present study demonstrates that fetal valves have much higher cellular densities than adult valves, associated with an increased cell proliferation-to-apoptosis ratio. VIC density was highest in the second trimester, &10-fold higher than in adult valves, and decreased progressively throughout gestation and postnatally. Fetal VIC proliferation indices were likewise greater than those of adult valves. Lincoln and colleagues²⁴ demonstrated decreased proliferation indices in chicken embryo valve primordia (calculated as the number of bromodeoxyuridine-positive nuclei/total nuclei), which supports the present finding that valve maturation is accompanied by decreased proliferation. We speculate that the higher rates of apoptosis in the late gestational period may reinforce the decreased cell proliferation at that interval to more rapidly and effectively arrest cuspal growth. The present study also demonstrates that valvular cell turnover is high during fetal development and continues at a low rate postnatally (Figure 2C) and that the total number of cells decreases substantially throughout life (Figure 2A), a fact that might account for slower or incomplete valve remodeling in older adults and predisposition to clinically important valve degeneration.

Activated Myofibroblasts Mediate Collagen Metabolism During Valve Maturation
We also demonstrated VIC plasticity throughout gestational valve maturation, in contrast to the resting VIC population of normal adult valves and their predominantly fibroblast-like phenotype.⁸ We observed that a large number of fetal VICs exhibited an -SMA–positive phenotype attributed to myofibroblast-like cells (Figure 3). Diseased valves also contain numerous myofibroblast-like VICs that express -SMA, which suggests a role for myofibroblasts in valvular connective tissue remodeling.^7,8 Moreover, we found that fetal VICs had abundant SMemb (nonmuscle myosin produced by embryonic or activated cells), MMP-1/collagenase-1, and MMP-13/collagenase-3, which indicates an activated/immature phenotype compatible with matrix remodeling. Despite the well-established role played by MMPs in pathological processes, including myxomatous mitral valve degeneration⁶ and atherosclerosis,^10,15 their role in remodeling normal valves and in fetal valve development remains unknown. We found more MMP-1 and MMP-13 in fetal valves, particularly late in gestation, than in adult valves, whereas collagen content increased during valve maturation. This inverse relationship suggests a critical role for MMP-family collagenases in collagen metabolism in cardiac valve development. A significant increase in collagen fiber thickness and alignment occurred between childhood and adulthood, whereas collagenase expression was negligible; however, whether this is due to prolonged exposure to mechanical forces or reflects completion of cell- and enzyme-associated remodeling is unknown.

Activated Myofibroblasts Undergo Phenotypic Modulation at Birth and Become a Quiescent Fibroblast-Like Cell Type During Adulthood
Previous studies reported that altered mechanical forces^8,26 and injury²⁷ are associated with phenotypic changes of valvular cells. Of particular interest in the present study was the way in which valves responded to abrupt change in blood pressures and other hemodynamic conditions at birth after separation of the single fetal circulation into isolated pulmonary and systemic circulations. Therefore, we also focused on comparison between fetal and neonatal pulmonary and aortic valves. Blood pressure in the fetal aorta and pulmonary artery is approximately 50/15 mm Hg. At birth, pulmonary pressure falls to 30/15 mm Hg; however, arterial pressure increases to 70/40 mm Hg. We found that the change from fetal to neonatal circulation correlated with reduced VIC activation (&6%) in neonatal pulmonary valves versus aortic valves (&21%; Figure 4).²⁸ Whether these changes might also be stimulated by changes in local blood oxygen content is unknown. In children’s valves, the numbers of activated VIC were similar in both pulmonary and aortic locations, which suggests tissue adaptation to pulmonary and systemic circulations.

Endothelial Cells Express an Activated Phenotype Throughout Fetal Development, a Phenotype Distinct From Adult Valves
Observations suggest that valvular endothelium is unique compared with other types of endothelial cells (particularly aortic and cardiac microvascular endothelium).²⁹ The endothelium regulates vascular tone, inflammation, thrombosis, and vascular remodeling.³⁰ Structurally intact endothelial cells can respond to pathophysiological stimuli by adjusting their usual functions and by expressing newly acquired properties, a process termed "endothelial activation." Activated endothelial cells produce a variety of biologically active products, including cytokines, growth factors, proteolytic enzymes, and adhesion molecules. Normal endothelial function is characterized by a balance of these factors and the cells’ ability to respond appropriately to stimuli. The present study demonstrated physiological activation of endothelial cells that consistently expressed high levels of SMemb, MMP-1, MMP-13, ICAM-1, and VCAM-1 in fetal and children’s valves. In contrast, adult valves had negligible expression of these molecules (Figure 5).

Clinical and experimental evidence suggests that valve leaflet endothelium is protected from inflammatory and immune-mediated injury by reduced expression of adhesion molecules, including VCAM-1, which is abundant in cardiac microvascular endothelium of rejected orthotopic heart transplants and in pig hearts after transplantation into baboons.³¹ The present study demonstrates consistent ICAM-1 and VCAM-1 expression by human fetal VECs during mid to late gestation. Furthermore, ICAM-1 and VCAM-1 expression has been found in surgically removed diseased valves^32,33 and in vitro,³⁴ which suggests that other factors, such as hemodynamic forces, may contribute to protection from inflammatory cell infiltration. Nevertheless, understanding endothelial adhesion molecule function in vivo in the presence of complex blood flow patterns requires further investigation.

Study Implications
The present study provides a natural history of cell and matrix changes in valve development, maturation, and adaptation (summarized in Figure 8) and extends the paradigm, previously suggested by us, that cardiac valvular tissue can adapt to environmental conditions, particularly mechanical loading.⁸ Specifically, under equilibrium conditions, VICs are quiescent and ECM is well adapted. We hypothesize that when stimulated by mechanical loading, VICs become activated and mediate connective tissue remodeling to restore a normal stress profile in the tissue. When equilibrium is restored, cells return to quiescence. The present study demonstrates that fetal VIC activation occurs throughout development, analogous to the valve changes that occur in pathological conditions and after surgical substitution.^6,7 Thus, analogous molecular mechanisms likely direct both physiological and pathological interstitial cell activation. Furthermore, our characterization of human semilunar valve VIC and VEC phenotypes and dynamic ECM changes from fetus to adult also indicates potential markers of cell activation/maturation and matrix alterations (eg, MMP-1, MMP-13, embryonic myosin, VCAM-1, and ICAM-1). These molecules are overexpressed during fetal valve development, several valvular diseases, and surgical transplantation and in tissue-engineered valves^7–9 and therefore may represent targets for molecular imaging to noninvasively monitor biological processes during healing and remodeling. Moreover, the progressive age-associated decrease in cell number and active capacity for remodeling may be an important clue in understanding senile valve degeneration.

View larger version (19K): [in this window] [in a new window]   Figure 8. Schematic representation of natural history of cardiac valve remodeling by interstitial cells. Schematic summary of cardiac valve remodeling in fetal valves according to the findings of the present study. Activated/immature cell phenotypes and collagen remodeling in gestation were followed after birth by decreased cell activation and eventually, in adults, by quiescence. These cellular changes were accompanied by increased collagen fiber thickness and alignment. Thin lines represent thin, immature collagen fibers; thick lines represent thick, mature collagen fibers.

Conclusions
We demonstrated that fetal valves possess a dynamic/adaptive structure and contain activated/immature cells. Moreover, the valvular cells that were activated in utero undergo phenotypic changes at birth and gradually become quiescent, whereas collagen matures through increased fiber thickness and alignment. This suggests a progressive adaptation to the prevailing hemodynamic environment. Knowledge of cell and matrix changes during fetal valve maturation may aid in the development of therapy for valve disease and assist in the design and evaluation of tissue-engineered valves.^35–37

	Acknowledgments

 
This study was supported in part by a grant from the Donald W. Reynolds Foundation (to Peter Libby, MD, Cardiovascular Division, Department of Medicine, Brigham and Women’s Hospital). The authors thank Vincent M. Lok, BA, and Matthias Nahrendorf, MD (Center for Molecular Imaging Research, Massachusetts General Hospital), for assistance with quantitative image analyses.

Disclosures

None.

	Footnotes

 
Guest Editor for this article was William C. Roberts, MD.

	References

Top Abstract Introduction Methods Results Discussion References

 
1. Eisenberg LM, Markwald RR. Molecular regulation of atrioventricular valvuloseptal morphogenesis. Circ Res. 1995; 77: 1–6.[Free Full Text]

2. Schroeder JA, Jackson LF, Lee DC, Camenisch TD. Form and function of developing heart valves: coordination by extracellular matrix and growth factor signaling. J Mol Med. 2003; 81: 392–403.[CrossRef][Medline] [Order article via Infotrieve]

3. Hurle JM. Scanning and light microscope studies of the development of the chick embryo semilunar heart valves. Anat Embryol (Berl). 1979; 157: 69–80.[CrossRef][Medline] [Order article via Infotrieve]

4. Hurle JM, Colvee E, Blanco AM. Development of mouse semilunar valves. Anat Embryol (Berl). 1980; 160: 83–91.[CrossRef][Medline] [Order article via Infotrieve]

5. Hurle JM, Colvee E. Changes in the endothelial morphology of the developing semilunar heart valves: a TEM and SEM study in the chick. Anat Embryol (Berl). 1983; 167: 67–83.[CrossRef][Medline] [Order article via Infotrieve]

6. Rabkin E, Aikawa M, Stone JR, Libby P, Schoen FJ. Activated interstitial myofibroblasts express catabolic enzymes and mediate matrix remodeling in myxomatous heart valves. Circulation. 2001; 104: 2525–2532.[Abstract/Free Full Text]

7. Rabkin-Aikawa E, Aikawa M, Farber M, Kratz JR, Garcia-Cardena G, Kouchoukos NT, Mitchell MB, Jonas RA, Schoen FJ. Clinical pulmonary autograft valves: pathologic evidence of adaptive remodeling in the aortic site. J Thorac Cardiovasc Surg. 2004; 128: 552–561.[Abstract/Free Full Text]

8. Rabkin-Aikawa E, Farber M, Aikawa M, Schoen FJ. Dynamic and reversible changes of interstitial cell phenotype during remodeling of cardiac valves. J Heart Valve Dis. 2004; 13: 841–847.[Medline] [Order article via Infotrieve]

9. Rabkin E, Hoerstrup SP, Aikawa M, Schoen FJ. Evolution of cell phenotype and extracellular matrix in tissue-engineered heart valves during in-vitro maturation and in-vivo remodeling. J Heart Valve Dis. 2002; 11: 308–314.[Medline] [Order article via Infotrieve]

10. Aikawa M, Rabkin E, Okada Y, Voglic SJ, Clinton SK, Brinckerhoff CE, Sukhova GK, Libby P. Lipid lowering by diet reduces matrix metalloproteinase activity and increases collagen content of rabbit atheroma: a potential mechanism of lesion stabilization. Circulation. 1998; 97: 2433–2444.[Abstract/Free Full Text]

11. Schurch W, Seemayer TA, Gabbiani G. The myofibroblast: a quarter century after its discovery. Am J Surg Pathol. 1998; 22: 141–147.[CrossRef][Medline] [Order article via Infotrieve]

12. Aikawa M, Sivam PN, Kuro-o M, Kimura K, Nakahara K, Takewaki S, Ueda M, Yamaguchi H, Yazaki Y, Periasamy M. Human smooth muscle myosin heavy chain isoforms as molecular markers for vascular development and atherosclerosis. Circ Res. 1993; 73: 1000–1012.[Abstract/Free Full Text]

13. Hiss J, Hirshberg A, Dayan DF, Bubis JJ, Wolman M. Aging of wound healing in an experimental model in mice. Am J Forensic Med Pathol. 1988; 9: 310–312.[CrossRef][Medline] [Order article via Infotrieve]

14. Rich L, Whittaker P. Collagen and picrosirius red staining: a polarized light assessment of fibrillar hue and spatial distribution. Br J Morphol Sci. 2005; 22: 97–104.

15. Deguchi J, Aikawa E, Libby P, Vachon JR, Inada M, Krane SM, Whittaker P, Aikawa M. MMP-13/collagenase-3 deletion promotes collagen accumulation and organization in mouse atherosclerotic plaques. Circulation. 2005; 112: 2708–2715.[Abstract/Free Full Text]

16. Whittaker P, Kloner RA, Boughner DR, Pickering JG. Quantitative assessment of myocardial collagen with picrosirius red staining and circularly polarized light. Basic Res Cardiol. 1994; 89: 397–410.[CrossRef][Medline] [Order article via Infotrieve]

17. Whittaker P, Canham PB. Demonstration of quantitative fabric analysis of tendon collagen using two-dimensional polarized light microscopy. Matrix. 1991; 11: 56–62.[Medline] [Order article via Infotrieve]

18. Oldenbourg R, Mei G. New polarized light microscope with precision universal compensator. J Microsc. 1995; 180: 140–147.[Medline] [Order article via Infotrieve]

19. Batschelet E, Hillman D, Smolensky M, Halberg F. Angular-linear correlation coefficient for rhythmometry and circannually changing human birth rates at different geographic latitudes. Int J Chronobiol. 1973; 1: 183–202.[Medline] [Order article via Infotrieve]

20. Schoen FJ. Aortic valve structure-function correlations: role of elastic fibers no longer a stretch of the imagination. J Heart Valve Dis. 1997; 6: 1–6.[Medline] [Order article via Infotrieve]

21. Johnson P, Maxwell DJ, Tynan MJ, Allan LD. Intracardiac pressures in the human fetus. Heart. 2000; 84: 59–63.[Abstract/Free Full Text]

22. Maron BJ, Hutchins GM. The development of the semilunar valves in the human heart. Am J Pathol. 1974; 74: 331–344.[Medline] [Order article via Infotrieve]

23. Poelmann RE, Mikawa T, Gittenberger-de Groot AC. Neural crest cells in outflow tract septation of the embryonic chicken heart: differentiation and apoptosis. Dev Dyn. 1998; 212: 373–384.[CrossRef][Medline] [Order article via Infotrieve]

24. Lincoln J, Alfieri CM, Yutzey KE. Development of heart valve leaflets and supporting apparatus in chicken and mouse embryos. Dev Dyn. 2004; 230: 239–250.[CrossRef][Medline] [Order article via Infotrieve]

25. Fisher SA, Langille BL, Srivastava D. Apoptosis during cardiovascular development. Circ Res. 2000; 87: 856–864.[Abstract/Free Full Text]

26. Weston MW, Yoganathan AP. Biosynthetic activity in heart valve leaflets in response to in vitro flow environments. Ann Biomed Eng. 2001; 29: 752–763.[CrossRef][Medline] [Order article via Infotrieve]

27. Tamura K, Jones M, Yamada I, Ferrans VJ. Wound healing in the mitral valve. J Heart Valve Dis. 2000; 9: 53–63.[Medline] [Order article via Infotrieve]

28. Shepherd JV, Vanhoutte PM. The Human Cardiovascular System: Facts and Concepts. New York, NY: Raven Press; 1979: 270–274.

29. Butcher JT, Penrod AM, Garcia AJ, Nerem RM. Unique morphology and focal adhesion development of valvular endothelial cells in static and fluid flow environments. Arterioscler Thromb Vasc Biol. 2004; 24: 1429–1434.[Abstract/Free Full Text]

30. Schoen FJ. Blood vessels. In: Kumar V, Fausto N, Abbas A, eds. Robbins and Cotran Pathologic Basis of Disease. 7th ed. Philadelphia, Pa: WB Saunders; 2004: 511–554.

31. Mitchell RN, Jonas RA, Schoen FJ. Pathology of explanted cryopreserved allograft heart valves: comparison with aortic valves from orthotopic heart transplants. J Thorac Cardiovasc Surg. 1998; 115: 118–127.[Abstract/Free Full Text]

32. Muller AM, Cronen C, Kupferwasser LI, Oelert H, Muller KM, Kirkpatrick CJ. Expression of endothelial cell adhesion molecules on heart valves: up-regulation in degeneration as well as acute endocarditis. J Pathol. 2000; 191: 54–60.[CrossRef][Medline] [Order article via Infotrieve]

33. Ghaisas NK, Foley JB, O’Briain S, Crean P, Kelleher D, Walsh M. Adhesion molecules in nonrheumatic aortic valve disease: endothelial expression, serum levels and effects of valve replacement. J Am Coll Cardiol. 2000; 36: 2257–2262.[Abstract/Free Full Text]

34. Dvorin EL, Jacobson J, Roth SJ, Bischoff J. Human pulmonary valve endothelial cells express functional adhesion molecules for leukocytes. J Heart Valve Dis. 2003; 12: 617–624.[Medline] [Order article via Infotrieve]

35. Rabkin E, Schoen FJ. Cardiovascular tissue engineering. Cardiovasc Pathol. 2002; 11: 305–317.[CrossRef][Medline] [Order article via Infotrieve]

36. Rabkin-Aikawa E, Mayer JE Jr, Schoen FJ. Heart valve regeneration. Adv Biochem Eng Biotechnol. 2005; 94: 141–179.[Medline] [Order article via Infotrieve]

37. Mol A, Bouten CV, Baaijens FP, Zund G, Turina MI, Hoerstrup SP. Tissue engineering of semilunar heart valves: current status and future developments. J Heart Valve Dis. 2004; 13: 272–280.[Medline] [Order article via Infotrieve]

This article has been cited by other articles:

				K. Schenke-Layland, U. A. Stock, A. Nsair, J. Xie, E. Angelis, C. G. Fonseca, R. Larbig, A. Mahajan, K. Shivkumar, M. C. Fishbein, et al. Cardiomyopathy is associated with structural remodelling of heart valve extracellular matrix Eur. Heart J., September 2, 2009; 30(18): 2254 - 2265. [Abstract] [Full Text] [PDF]

				M. D. Combs and K. E. Yutzey Heart Valve Development: Regulatory Networks in Development and Disease Circ. Res., August 28, 2009; 105(5): 408 - 421. [Abstract] [Full Text] [PDF]

				J. P. Dal-Bianco, E. Aikawa, J. Bischoff, J. L. Guerrero, M. D. Handschumacher, S. Sullivan, B. Johnson, J. S. Titus, Y. Iwamoto, J. Wylie-Sears, et al. Active Adaptation of the Tethered Mitral Valve: Insights Into a Compensatory Mechanism for Functional Mitral Regurgitation Circulation, July 28, 2009; 120(4): 334 - 342. [Abstract] [Full Text] [PDF]

				E. Aikawa, M. Aikawa, P. Libby, J.-L. Figueiredo, G. Rusanescu, Y. Iwamoto, D. Fukuda, R. H. Kohler, G.-P. Shi, F. A. Jaffer, et al. Arterial and Aortic Valve Calcification Abolished by Elastolytic Cathepsin S Deficiency in Chronic Renal Disease Circulation, April 7, 2009; 119(13): 1785 - 1794. [Abstract] [Full Text] [PDF]

				P. Sucosky, K. Balachandran, A. Elhammali, H. Jo, and A. P. Yoganathan Altered Shear Stress Stimulates Upregulation of Endothelial VCAM-1 and ICAM-1 in a BMP-4- and TGF-{beta}1-Dependent Pathway Arterioscler Thromb Vasc Biol, February 1, 2009; 29(2): 254 - 260. [Abstract] [Full Text] [PDF]

				A. Balguid, A. Mol, M. A.A. van Vlimmeren, F. P.T. Baaijens, and C. V.C. Bouten Hypoxia Induces Near-Native Mechanical Properties in Engineered Heart Valve Tissue Circulation, January 20, 2009; 119(2): 290 - 297. [Abstract] [Full Text] [PDF]

				F. J. Schoen Evolving Concepts of Cardiac Valve Dynamics: The Continuum of Development, Functional Structure, Pathobiology, and Tissue Engineering Circulation, October 28, 2008; 118(18): 1864 - 1880. [Abstract] [Full Text] [PDF]

				M. Chaput, M. D. Handschumacher, F. Tournoux, L. Hua, J. L. Guerrero, G. J. Vlahakes, and R. A. Levine Mitral Leaflet Adaptation to Ventricular Remodeling: Occurrence and Adequacy in Patients With Functional Mitral Regurgitation Circulation, August 19, 2008; 118(8): 845 - 852. [Abstract] [Full Text] [PDF]

				R. B. Hinton Jr., C. M. Alfieri, S. A. Witt, B. J. Glascock, P. R. Khoury, D. W. Benson, and K. E. Yutzey Mouse heart valve structure and function: echocardiographic and morphometric analyses from the fetus through the aged adult Am J Physiol Heart Circ Physiol, June 1, 2008; 294(6): H2480 - H2488. [Abstract] [Full Text] [PDF]

				K. B. Donnelly Cardiac Valvular Pathology: Comparative Pathology and Animal Models of Acquired Cardiac Valvular Diseases Toxicol Pathol, February 1, 2008; 36(2): 204 - 217. [Abstract] [Full Text] [PDF]

				R. F. Padera Jr. and F. J. Schoen Pathology of Cardiac Surgery Card. Surg. Adult, January 1, 2008; 3(2008): 111 - 178. [Full Text]

				J. E. Mayer Tissue Engineering for Cardiac Valve Surgery Card. Surg. Adult, January 1, 2008; 3(2008): 1649 - 1656. [Full Text]

				E. Aikawa, M. Nahrendorf, J.-L. Figueiredo, F. K. Swirski, T. Shtatland, R. H. Kohler, F. A. Jaffer, M. Aikawa, and R. Weissleder Osteogenesis Associates With Inflammation in Early-Stage Atherosclerosis Evaluated by Molecular Imaging In Vivo Circulation, December 11, 2007; 116(24): 2841 - 2850. [Abstract] [Full Text] [PDF]

				A. C. Liu, V. R. Joag, and A. I. Gotlieb The Emerging Role of Valve Interstitial Cell Phenotypes in Regulating Heart Valve Pathobiology Am. J. Pathol., November 1, 2007; 171(5): 1407 - 1418. [Abstract] [Full Text] [PDF]

				A. H Chester and P. M Taylor Molecular and functional characteristics of heart-valve interstitial cells Phil Trans R Soc B, August 29, 2007; 362(1484): 1437 - 1443. [Abstract] [Full Text] [PDF]

				E. Aikawa, M. Nahrendorf, D. Sosnovik, V. M. Lok, F. A. Jaffer, M. Aikawa, and R. Weissleder Multimodality Molecular Imaging Identifies Proteolytic and Osteogenic Activities in Early Aortic Valve Disease Circulation, January 23, 2007; 115(3): 377 - 386. [Abstract] [Full Text] [PDF]

				S. Paruchuri, J.-H. Yang, E. Aikawa, J. M. Melero-Martin, Z. A. Khan, S. Loukogeorgakis, F. J. Schoen, and J. Bischoff Human Pulmonary Valve Progenitor Cells Exhibit Endothelial/Mesenchymal Plasticity in Response to Vascular Endothelial Growth Factor-A and Transforming Growth Factor-{beta}2 Circ. Res., October 13, 2006; 99(8): 861 - 869. [Abstract] [Full Text] [PDF]

				M. Nahrendorf, F. A. Jaffer, K. A. Kelly, D. E. Sosnovik, E. Aikawa, P. Libby, and R. Weissleder Noninvasive Vascular Cell Adhesion Molecule-1 Imaging Identifies Inflammatory Activation of Cells in Atherosclerosis Circulation, October 3, 2006; 114(14): 1504 - 1511. [Abstract] [Full Text] [PDF]

				R. B. Hinton Jr, J. Lincoln, G. H. Deutsch, H. Osinska, P. B. Manning, D. W. Benson, and K. E. Yutzey Extracellular Matrix Remodeling and Organization in Developing and Diseased Aortic Valves Circ. Res., June 9, 2006; 98(11): 1431 - 1438. [Abstract] [Full Text] [PDF]

This Article Abstract Full Text (PDF) Alert me when this article is cited Alert me if a correction is posted Citation Map Services Email this article to a friend Similar articles in this journal Similar articles in PubMed Alert me to new issues of the journal Download to citation manager Request Permissions Citing Articles Citing Articles via HighWire Citing Articles via Google Scholar Google Scholar Articles by Aikawa, E. Articles by Schoen, F. J. Search for Related Content PubMed PubMed Citation Articles by Aikawa, E. Articles by Schoen, F. J. Related Collections Valvular heart disease CV surgery: valvular disease